Architecture-Dependent Stabilization of Polyelectrolyte Complexes

Article pubs.acs.org/Macromolecules

Architecture-Dependent Stabilization of Polyelectrolyte Complexes between Polyanions and Cationic Triblock Terpolymer Micelles Jennifer E. Laaser,† Elise Lohmann,† Yaming Jiang,‡ Theresa M. Reineke,† and Timothy P. Lodge*,†,‡ †

Department of Chemistry and ‡Department of Chemical Engineering & Materials Science, University of Minnesota, Minneapolis, Minnesota 55455, United States S Supporting Information *

ABSTRACT: We investigate the complexation of poly(styrenesulfonate) with micelles containing both cationic and hydrophilic blocks in their coronas. Five distinct micelles were prepared by self-assembly, using D+S, OS, OD+S, D+OS, and mixtures of D+S and OS block polymers, where the hydrophobic S blocks (poly(styrene)) form the micelle cores and the cationic D+ blocks (poly(dimethylaminoethyl methacrylate)) and hydrophilic, nonionic O blocks (poly(oligo(ethylene glycol) methyl ether methacrylate)) form the coronas. Turbidimetric titration and dynamic light scattering measurements on complexes with short poly(styrenesulfonate) chains (M ≈ 1 kg/mol) that can equilibrate quickly reveal that the intrinsic colloidal stability of the complexes is determined by the identity of the outermost block of the micelle corona and that architectures with a nonionic solvating outer block promote the formation of soluble single-micelle complexes even when the complexes are fully neutralized. Although complexes with longer poly(styrenesulfonate) chains (M ≈ 30 kg/mol) are kinetically trapped in aggregates for all cationcontaining micelle architectures, studies at high ionic strength show that inclusion of the outer hydrophilic block can successfully limit the size of the complexes and inhibit overall phase separation of neutralized complexes. Finally, the molecular weight dependence of the aggregation process for complexes of the OD+S architecture demonstrates that bridging is the predominant mechanism for aggregation and that careful selection of the polymer architecture and molecular weight can provide a useful strategy for controlling the structure and colloidal stability of hierarchical complexes.

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INTRODUCTION Complexation of oppositely charged polyelectrolytes is of intense interest, both as a fundamental problem in polymer physics and as a key process underlying materials and applications ranging from surface coatings,1 flocculants,2,3 and separation membranes4 to biocompatible materials,1 hydrogels,5−7 and drug and nucleic acid delivery vehicles.8−13 A common feature in many systems forming polyelectrolyte complexes is insolubility of the complexes near the 1:1 (or stoichiometric) charge ratio.14−16 While macroscopic phase separation in these systems is desirable for some applications (e.g., wastewater treatment or membranes), it is problematic in many others (e.g., nucleic acid or protein delivery) that require the polyelectrolyte complexes to form small, soluble structures that can be used to transport cargo. Controlling the complex structure and solubility is thus critical for optimizing the material properties necessary for different applications. Numerous efforts have been made to improve the colloidal stability of polyelectrolyte complexes via addition of solvating, nonionic blocks to one or both of the polyelectrolytes that make up the complex. Below a critical salt concentration, polyelectrolyte complexes of these hydrophilic diblock polyelectrolytes typically form so-called “complex coacervate core” micelles, in which the hydrophilic nonionic chains form a soluble shell around an insoluble core of charge-neutral polyelectrolyte complex.17−22 The structure and stability of © XXXX American Chemical Society

these micelles depend on the energetic balance of the surface tension between the neutralized polyelectrolyte region and the surrounding water, with the stretching of the solvating chains in the micelle corona.11,23,24 Triblock architectures with a hydrophilic chain sandwiched between two charged ends can also give rise to responsive hydrogel networks of complex coacervate core micelles connected by bridging nonionic, hydrophilic chains.5,6 The polymer architecture, along with solution conditions such as pH and ionic strength,6,18,19,22 thus provides a way to control the structure, solubility, and materials properties of polyelectrolyte complexes for a wide range of applications. In nucleic acid delivery systems, in which a polyelectrolyte complex is formed between a synthetic polycation and DNA or RNA, incorporating carefully chosen hydrophilic blocks into the polycation architecture can also reduce unwanted side effects such as capillary blockage and immune recognition/macrophage uptake, and thus improve the circulation time of the complexes, promote tissue penetration and cellular uptake, and prevent loss of cargo by inhibiting polyion exchange with other charged species present in the blood such as serum proteins.11,12,23,25 Received: June 30, 2016 Revised: August 15, 2016

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DOI: 10.1021/acs.macromol.6b01408 Macromolecules XXXX, XXX, XXX−XXX

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Macromolecules As a general platform for such applications, polyelectrolyte micelles with a hydrophobic core and a charged corona carry several potential advantages, such as the ability to codeliver hydrophobic cargo sequestered in the micelle core simultaneously with the polyelectrolyte complexed in the corona. For example, this approach has been successfully used to codeliver chemotherapy drugs alongside small interfering RNA (siRNA) for cancer treatment.8,26 However, less is known about how polymer architecture may be used to control the structure, solubility, and stability of these complexes. When complexed with an oppositely charged polyelectrolyte, diblock polyelectrolyte micelles typically form overcharged multimicelle aggregates that are solubilized by chains of the polyelectrolyte present in excess when prepared far from the stoichiometric charge ratio, but that precipitate from solution when fully neutralized.27−33 Complexation of polyelectrolyte micelles with an oppositely charged polyelectrolyte containing a hydrophilic block, on the other hand, can lead to formation of layered micelles solvated by the hydrophilic chains.34−36 A number of studies have shown that micelle-forming triblock polymers containing a hydrophilic nonionic block alongside the charged block and the core-forming hydrophobic block can help solubilize complexes of these micelles with nucleic acids for delivery applications.10,26,37,38 However, these studies have focused primarily on the application-specific performance of the materials and less on understanding the effects of interpolyelectrolyte interactions on phase behavior and stability of the complexes. In this paper, we investigate the complexation of cationic polymeric micelles with an oppositely charged homopolymer as a function of micelle architecture and polyanion molecular weight, in order to elucidate the design rules for soluble micelle−polyelectrolyte systems and bring insight into the mechanism of the complexation process. We systematically vary the corona architecture by preparing well-defined block polymers with different block chemistries and sequences. The block polymers are composed of hydrophobic S blocks (poly(styrene), PS), cationic D+ blocks (protonated poly(dimethylaminoethyl methacrylate), PDMAEMA), and hydrophilic, nonionic O blocks (poly(oligo(ethylene glycol) methyl ether methacrylate), POEGMA). PDMAEMA-containing polymers and their complexes with poly(styrenesulfonate) (PSS) are a useful model system for applications such as nucleic acid delivery and have been well-characterized in previous work,9,13,33,39−42 making them an attractive system on which to base studies of more complex architectures. As shown in Figure 1, four block polymers are used to prepare five different micelle architectures in this work. These architectures include micelles formed directly from individual D+S, OS, OD+S, and D+OS block polymers, which have layers of cationic and hydrophilic, nonionic units in the coronas, as well as micelles formed from a mixture of the D+S and OS block polymers, which contain a mixture of charged and uncharged chains in the micelle corona. The polymers are designed such that the cationic and hydrophilic, nonionic blocks have similar molecular weights, facilitating direct elucidation of architecture-dependent effects on the complexation process. The results demonstrate that the intrinsic colloidal stability of the complexes is controlled primarily by the outermost corona block and provide new insight into both the mechanisms of complexation and how polymer architecture can be used to control the structure and stability of micelle−polyelectrolyte complexes.

Figure 1. Schematic of micelle architectures used in this work. Styrene constitutes the hydrophobic (S) block of the polymers and is buried in the core of the resulting micelles, while the coronas are made up of dimethylaminoethyl methacrylate (D), which is hydrophilic and cationic in the acidic solutions used in this work (denoted D+), and oligo(ethylene glycol) methyl ether methacrylate, which is also hydrophilic but carries no charge (O).

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MATERIALS AND METHODS

Materials. Styrene, N,N-dimethylaminoethyl methacrylate (DMAEMA), oligo(ethylene glycol) methyl ether methacrylate (OEGMA, Mn = 500 g/mol), 4,4′-azobis(4-cyanovaleric acid) (ACVA), azobis(isobutyronitrile) (AIBN), and 4-cyano-4-[(dodecylsulfanylthiocarbonyl)sulfanyl]pentanoic acid (CDTPA) were purchased from Sigma-Aldrich. Styrene, DMAEMA, and OEGMA were filtered through activated neutral alumina to remove inhibitor before use. AIBN was recrystallized once from methanol, and ACVA and CDTPA were used as received. Poly(styrenesulfonate) (PSS) was purchased from Polymer Standards Service (Mainz, Germany). The PSS samples used in this work had weight-average molecular weights (Mw) of 1.1 kg/mol (PSS-1) and 29 kg/mol (PSS-30), both with quoted dispersities less than 1.2 and degrees of sulfonation greater than 95%. Polymer Synthesis. The block polymers were synthesized by multistep reversible addition−fragmentation chain transfer (RAFT) polymerization, as has been described previously.41 Full synthetic details for all polymers are provided in the Supporting Information. Briefly, for each block, the monomer, chain transfer agent (CDTPA or previously synthesized polymeric macro-chain transfer agent), and initiator were dissolved in toluene, degassed via three freeze−pump− thaw cycles, and polymerized at 80 °C until 1H nuclear magnetic resonance (NMR) spectra of the reaction mixture indicated that the target conversion had been reached. The reaction was then quenched by cooling to room temperature and exposure to atmosphere. Unreacted monomer was removed by precipitation (for polymerizations of styrene or DMAEMA) or dialysis (for polymerizations of OEGMA), and the resulting polymers were then dried and characterized by NMR and size exclusion chromatography (SEC). NMR measurements were made on a 500 MHz Bruker HD-500 spectrometer equipped with a cryoprobe. SEC measurements were carried out on a system equipped with an Agilent 1260 pump, multiangle light-scattering detector (Wyatt Dawn DSP-F), and refractive index detector (Wyatt Optilab DSP) for absolute determination of the polymer molecular weights. All SEC measurements were made at room temperature using THF with 1% tetramethylethylenediamine as the eluent. Literature values of 0.186 and 0.086 mL/g were used for the refractive index increments (dn/dc) of poly(styrene) and poly(dimethylaminoethyl methacrylate), respectively,41,39 while that of POEGMA was measured by direct injection of a series of POEGMA solutions through the refractive index detector, yielding a value of 0.075 mL/g. The final synthesized polymers were DS(30-8) (Đ = 1.09), OS(31-25) (Đ = 1.19), DOS(30-28-12) (Đ = 1.17), and ODS(31-26-7) (Đ = 1.23), where D indicates a DMAEMA block, O indicates an OEGMA block, S indicates a styrene block, and the numbers in parentheses denote the number-average molecular weights of each block. Full characterization data for these polymers and the intermediate macro-chain transfer agents are provided in Table 1. B

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that the weight fraction of OEGMA in solution matched the weight fraction of DMAEMA in the DS(30-8) samples). PSS solutions were prepared in the same buffers by direct dissolution in the leftover dialysate to a final concentration of sulfonate units of 1.2 mM. These concentrations are consistent with the dilute solution conditions used in previous work on complexes of polyelectrolyte micelles.28,30,32,33 Turbidimetric Titrations. Turbidimetric titrations were carried out using a home-built light transmission apparatus, as described previously.33 For each titration, 1.250 mL of the starting solution was placed in a vial with a stir bar. The oppositely charged polyelectrolyte was then added dropwise with a micropipet while stirring. After each 50 μL addition, the solution was allowed to equilibrate for 30 s, after which the total transmitted power of a 632 nm HeNe laser was measured on a Spex Industries laser power meter. To facilitate probing a large range of charge ratios with minimal sample volume, 500 μL of the mixed solution was removed after every tenth 50 μL addition. Formation of heterogeneous precipitates was assessed by visual inspection after every step. Dynamic Light Scattering. Samples for dynamic light scattering were prepared by a titration procedure similar to that used for the turbidimetric titration experiments. To minimize dust, the micelle and PSS stock solutions were filtered through 0.2 μm filters before use, and the glass vials and stir bars used for sample mixing were rinsed with filtered solvent before use. Prepared samples were then transferred directly into clean DLS sample tubes without further filtration. Dynamic light scattering measurements were made on a Brookhaven Instruments BI-200SM multiangle light scattering instrument equipped with a 637 nm laser diode and avalanche photodiode detector. Autocorrelation functions were acquired for time delays from 1 μs to 1 s at room temperature (23 °C) and a scattering angle of 90°, with data averaged for 30 min. Decay time distributions were then extracted using the REPES algorithm, and the resulting distributions were converted to hydrodynamic radius (Rh) distributions via the Stokes−Einstein relation using viscosities and refractive indices for equivalent ionic strength sodium chloride solutions. Finally, peak positions were extracted by fitting to a log-normal distribution.

Table 1. Summary of Polymer Characterization Data polymer

wDMAEMA

wOEGMA

wS

D(30) DS(30-8)

1 0.78

0 0

0 0.22

DO(30-28)

0.51

0.49

0

DOS(30-28-12)

0.42

0.45

0.13

O(31) OS(31-25)

0 0

1 0.56

0 0.44

OD(31-26)

0.46

0.54

0

ODS(31-26-7)

0.40

0.48

0.12

Mn (kg/mol) 30 46 38 68 58 82 71 31 57 55 57 57 65 64

(SEC) (NMR) (SEC) (NMR) (SEC) (NMR) (SEC) (NMR) (SEC) (NMR) (SEC) (NMR)

Đ 1.08 1.09 1.12 1.17 1.19 1.19 1.14 1.23

Micelle Formation. Micelles were formed from the synthesized block polymers by a cosolvent addition process, as described previously.41 Micelle stock solutions were prepared from each of the four block polymers alone as well as from a mixture of DS(30-8) and OS(31-25) (denoted DS(30-8)/OS(31-25)) containing equal amounts of DMAEMA and OEGMA by weight. Each block polymer or polymer mixture was dissolved in dimethylformamide (DMF) at a concentration of 10 mg/mL. An equal volume of 100 mM TRIS buffer at pH 7.25 was then added dropwise while stirring, followed by a further 2-fold dilution with TRIS buffer. The resulting micelle stock solutions were then dialyzed against 20 mM acetate buffer at pH 4.5 with a total ionic strength of either 10 or 500 mM set by addition of sodium chloride. Following dialysis, the solutions were diluted to a total concentration of 1.2 mM DMAEMA (for DS(30-8), ODS(31-267), DOS(30-28-12), and DS(30-8)/OS(31-25) micelles) or 0.39 mM OEGMA (for OS(31-25) micelles; this concentration was chosen so

Figure 2. Turbidimetric titrations of (a, b) DS(30-8) and (c, d) ODS(31-26-7) with (a, c) PSS-1 and (b, d) PSS-30 in acetate buffer at pH 4.5 and 10 mM ionic strength. Transmission is reported as a function of xamine, or the fraction of all charges in the system coming from protonated DMAEMA groups. C

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Figure 3. Summary of turbidimetric titration end points for all five micelle architectures with (a) PSS-1 and (b) PSS-30 in acetate buffer at pH 4.5 and 10 mM ionic strength. From lightest to darkest, blue bars indicate the regions between the 90, 85, and 80% transmission end points, as illustrated in the Supporting Information (Figure S11).

Figure 4. Hydrodynamic radius distributions for (a) DS(30-8) and (d) ODS(31-26-7) micelles complexed with (b, e) PSS-1 and (c, f) PSS-30 in acetate buffer at pH 4.5 and 10 mM ionic strength. Samples with excess PSS were prepared at xamine = 0.29, while samples with excess micelles were prepared at xamine = 0.71. The flat line in (b) indicates that the complexes precipitated and could not be characterized by DLS. Zeta Potentials. Zeta potentials were measured on a Zetasizer Nano ZS from Malvern Instruments. Samples were prepared as for the DLS measurements and were transferred to disposable folded capillary cells for measurement. Electrophoretic mobilities were measured using the Malvern M3-PALS method and converted to zeta potentials via the Smoluchowski equation. The reported values were taken from the average of three measurements. Samples typically had conductivities around 1 mS/cm. Cryogenic Transmission Electron Microscopy. Samples for cryogenic transmission electron microscopy (cryoTEM) were prepared using a FEI Vitrobot Mark III. For each sample, approximately 3 μL of solution was deposited on a 300 mesh lacey carbon/Formvar-coated copper TEM grid (Ted Pella). The sample was blotted for 5 s, drained for 1 s, and vitrified by plunging into liquid ethane. Samples were stored under liquid nitrogen and transferred to a single-tilt cryo holder for imaging. TEM images were acquired on a

FEI Tecnai G2 Spirit BioTWIN electron microscope operating at an accelerating voltage of 120 kV and equipped with an Eagle 4 megapixel CCD camera; images were acquired at a slight underfocus for adequate contrast. Complexes were typically vitrified within 1 week of preparation and in all cases during the window in which DLS indicated that the initial size distribution remained stable.

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RESULTS Turbidimetric Titrations. Turbidimetric titration curves for DS(30-8) and ODS(31-26-7) micelles complexed with PSS1 and PSS-30 at low ionic strength are presented in Figure 2. Titration curves for the other three polymer systems are provided in the Supporting Information (Figures S12−S14), but we focus here on the PDMAEMA-b-PS (DS) and POEGMA-b-PDMAEMA-b-PS (ODS) architectures because D

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Macromolecules Table 2. Zeta Potentials (in mV) of Micelles and Complexes at 10 mM Ionic Strength complexes with PSS-1 polymer DS(30-8) OS(31-25) ODS(31-26-7) DOS(30-28-12) DS(30-8)/OS(30-8) PSS alone a

micelles 28 −2 22 36 34

± ± ± ± ±

3a 0 1 1 2

xamine = 0.29

complexes with PSS-30

xamine = 0.71 22 −4 17 31 32

−7 ± 2 −3 ± 0

−15.5 ± 3.3

± ± ± ± ±

5 0 1 2 3

xamine = 0.29 −37 −6 −7 −16 −25

± ± ± ± ±

xamine = 0.71

1 1 1 1 1

37 −16 14 24 33

± ± ± ± ±

2 1 0 2 2

−9.9 ± 4.5

Reported uncertainties are the standard deviation of three consecutive measurements.

were found to be asymmetric when titrated with PSS-1. One particularly important observation highlighted in Figure 3 is that the ODS(31-26-7) micelles were the only system that exhibited precipitate formation and decreased transmission with PSS-30 but not with PSS-1. This observation brings insight into the mechanism of complexation and aggregation in these micelle−polyelectrolyte complexes, as will be discussed in more detail below. Dynamic Light Scattering. Hydrodynamic radius distributions for complexes of DS(30-8) and ODS(31-26-7) micelles with PSS-1 and PSS-30 at low ionic strength are presented in Figure 4. For both micelle architectures, complexes with PSS-30 exhibited bimodal size distributions that were stable for more than one month, again consistent with previous results on complexation of PDMAEMA-b-PS micelles.33 The soluble complexes with PSS-1, on the other hand, exhibited only monomodal size distributions with radii close to or slightly smaller than those of the uncomplexed micelles. As in the turbidimetric titration experiments, similar trends were observed in dynamic light scattering measurements on complexes of the other three micelle architectures. Hydrodynamic radius distributions for complexes of the OS(31-25), DOS(30-28-12), and DS(30-8)/OS(31-25) micelles with PSS1 and PSS-30 are presented in the Supporting Information

they provide clear examples of the different types of titration behavior observed as a function of polymer architecture and molecular weight. As shown in Figure 2, the titration curves depended strongly on both the micelle architecture and the polyanion molecular weight. When complexed with the longer PSS (PSS-30, Figures 2b and 2d), the titration curves were qualitatively the same for both micelle architectures and were similar to those previously observed for PDMAEMA-b-PS micelles.33 For both the forward and reverse titrations, the titration curves were relatively symmetric. The transmission decreased slowly until the charge ratio was close to the stoichiometric point (xamine = 0.5), near which the transmission dropped rapidly and an insoluble precipitate formed. When complexed with the short PSS oligomer (PSS-1), however, the shape of the titration curves was no longer the same for both micelle architectures and was significantly different from those observed in the titrations with PSS-30. For complexes with ODS(31-26-7), the solution remained clear and homogeneous, and the transmission remained steady at its initial value throughout the titration. For complexes with DS(30-8), the transmission did decrease and a precipitate was formed, but unlike the titrations with PSS-30, the curves for the forward and reverse titrations with PSS-1 changed shape and were no longer symmetric. When the micelles were added to the PSS, the transmission began to drop immediately, and an insoluble precipitate was observed as early as xamine = 0.095. The transmission dropped steadily until the stoichiometric point, after which it began to increase in a similar manner. Although the insoluble precipitate never completely redissolved, this increase in transmission did correspond to a decrease in the apparent particle size, and by the end of the titration, few precipitate particles remained visible. When the PSS was added to the micelles, on the other hand, the titration curve had a similar shape to those observed with the longer PSS, with a sharp drop in transmission and precipitate formation near the stoichiometric point. However, unlike the titrations with the longer PSS, the drop in transmission occurred almost exactly at the stoichiometric point (at xamine = 0.49 for PSS-1 added to DS(30-8)) rather than somewhat before (e.g., at xamine = 0.57 for PSS-30 added to DS(30-8)). As summarized in Figure 3 and Figures S12−S15, titration curves for the other three micelle architectures exhibited similar features when complexed with PSS-1 and PSS-30 at low ionic strength. As in the titrations of ODS(31-26-7) micelles with PSS-1, OS(31-25) micelles neither precipitated nor exhibited a measurable decrease in solution transmission when mixed with either length of PSS. DOS(30-28-12) and DS(30-8)/OS(3125) micelles, on the other hand, exhibited essentially the same titration behavior as the DS(30-8) micelles for both molecular weights of PSS. The titration curves were relatively symmetric for both the forward and reverse titrations with PSS-30 but

Figure 5. Temporal evolution of peak positions in dynamic light scattering data for all five micelle architectures complexed with (a, c) PSS-1 and (b, d) PSS-30 in acetate buffer at pH 4.5 and 10 mM ionic strength at (a, b) xamine = 0.29 and (c, d) xamine = 0.71. Samples that precipitated immediately upon preparation are omitted from the plot. E

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Figure 6. Hydrodynamic radius distributions for complexes of (a) DS(30-8) and (b) ODS(31-26-7) with PSS-30 in acetate buffer at pH 4.5 and 500 mM ionic strength.

impossible to characterize because they precipitated. Interestingly, complexes of ODS(31-26-7) with an excess of PSS-1, which was the only DMAEMA-containing micelle system to remain soluble with PSS-1 in excess, had a zeta potential of only −3 mV, very close to that of the DMAEMA-free OS(31-25) micelles. Cryogenic Transmission Electron Microscopy. Cryogenic transmission electron micrographs of ODS(31-26-7) micelles and their complexes with PSS-1 and PSS-30 are presented in Figure 7. While the uncomplexed micelle coronas were invisible in the cryoTEM images, the polystyrene cores, shown in Figure 7a, were generally round and uniform in size and were distributed on a hexagonal lattice resulting from packing of the spherical micelles during sample blotting and vitrification.43,44 Similar sizes and shapes were observed for the other four micelle systems, as shown in the Supporting Information (Figure S9). Complexes of ODS(31-26-7) with PSS-1 appeared to form well-separated single-micelle complexes that are similar in size to the uncomplexed micelles at xamine = 0.71, but are 2−3 times larger at xamine = 0.29, consistent with formation of a higher contrast polyelectrolyte complex shell around the micelle core.33 The complexes with PSS-30, on the other hand, appeared to form mixtures of single micelles with multimicelle aggregates, as has previously been observed for the PDMAEMA-b-PS system.33 The isolated single micelle species in these samples were similar in size to the uncomplexed micelles, while the particles making up the multimicelle aggregates were significantly larger. The particles making up the aggregates appeared larger and more homogeneous in the presence of excess PSS (xamine = 0.29) than in the presence of excess micelles (xamine = 0.71).

(Figures S16−S18), and their peak positions are summarized in Figure 5. As is evident from these figures, complexes of the DOS(30-28-12) and DS(30-8)/OS(31-25) micelle architectures formed similar size distributions to those observed for complexes of the DS(30-8) micelles, forming insoluble precipitates or soluble monomodal distributions when complexed with PSS-1 at xamine = 0.29 and 0.71, respectively, and stable, bimodal distributions when complexed with PSS-30 at either charge ratio. While the hydrodynamic radius distributions were stable for at least 4 weeks at low ionic strength (10 mM), the size of complexes formed at high ionic strength (500 mM) changed significantly in less than 1 week, as shown in Figure 6. For complexes of DS(30-8) with PSS-30 at high ionic strength, complexes at both charge ratios initially formed bimodal distributions comparable to those observed at low ionic strength. Complexes formed with the PSS in excess (xamine = 0.29) then grew larger and eventually precipitated, as has been observed previously,33 while those with the micelles in excess (xamine = 0.71) rearranged to a monomodal distribution with a size slightly smaller than that of the uncomplexed micelles. Complexes of ODS(31-26-7) exhibited similar behavior with the micelles in excess, rearranging to a monomodal distribution with a radius slightly smaller than that of the uncomplexed micelles in under a week. With the PSS in excess, however, the hydrodynamic radius distribution of complexes of ODS(31-267) remained relatively steady over the course of the experiment and, unlike the complexes of DS(30-8), remained soluble. Notably, while the initial sizes of the complexes of DS(30-8) with PSS-30 were similar at both 10 and 500 mM ionic strengths, the complexes of ODS(31-26-7) were significantly smaller when formed at 500 mM ionic strength than when formed at 10 mM ionic strength. Zeta Potentials. Zeta potentials were measured for complexes and micelles of all five micelle architectures at low ionic strength, as summarized in Table 2. There are several important trends evident in these data. First, while the OS(3125) micelles had zeta potentials very close to 0, at −2 mV, all DMAEMA-containing micelles had positive zeta potentials between 20 and 40 mV. After complexation with PSS-30, the zeta potentials for complexes formed with PSS in excess (xamine = 0.29) became negative, while those for complexes formed with the micelles in excess (xamine = 0.71) remained positive. For complexes formed with PSS-1, complexes formed with the micelles in excess similarly had positive zeta potentials, while those formed with the PSS in excess were in most cases

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DISCUSSION The goal of this paper is to investigate how corona architecture determines the structure and stability of micelle−polyelectrolyte complexes. The following discussion addresses three separate aspects of this problem: first, how block sequence and corona architecture affect the colloidal stability of micelle− polyelectrolyte complexes; second, how the polymer architecture that has the most pronounced effect on the solubility affects the structure of the complexes and the complexation mechanism; third, how corona architecture affects the kinetic trapping and annealing of micelle−polyelectrolyte complexes, and the implications of these results for the design of polyelectrolyte micelles for diverse applications. F

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DMAEMA-containing micelles. Additionally, cryoTEM images of the complexes formed with excess PSS revealed an increase in the size of the dense region surrounding the micelle core, consistent with polyelectrolyte complex formation, and the zeta potential of the complexes was significantly lower than that of the micelles alone, indicating neutralization of the positively charged DMAEMA region. In fact, the zeta potentials of the complexes of ODS(31-26-7) were low enough (